I am purifying a protein (47 kDa) that is fused onto an MBP (42 kDa). The PIs for both proteins are 6.48 and 4.9 respectively. After purifying it using a dextrose column, I have buckets of fused proteins (up to 30 mg/ml). I dialysed my proteins into a reaction buffer to rid of the maltose and to optimise the cleavage of the tag (20 mM Tris, 50 mM NaCl and 1 mM CaCl2 pH 7.5). Consequently, I tried cleaving it at small scale (10mg of protein with 100ug of TEV protease with 0.005% SDS) for 6 hours.
After cleavage, I dialysed my samples into a buffer (20 mM Tris, 50 mM NaCl and 1 mM CaCl2 pH 6.4) for anionic exchange. Theoretically, my proteins should be neutral and only other proteins would bind onto the anionic exchanger. The picture of the gel shows the flowthrough and the elution of my proteins. Lanes 1-3 show the elution of my protein while lanes 4-9 shows the elution of other proteins. So far, the bands corresponds to the size of my protein. However, whan I mass-spec my the bands on this gel, the results show that the bands on lane 3 is mainly MBP. the concentration of the flowthrough is 1.63 mg/ml.
This is my first time using MBP and purifying this protein. I can't think of where my protein might have disappeared. There were no signs of precipitation in the solution. Any ideas?
In the future I will be optimising the expression of my protein. e.g. inducing at room temperature overnight, adding glycerol in my buffers and conducting my experiments at 4C. I would also run my samples through another dextrose column after cleavage to rid of the MBP although I am not sure as to how this would improve my technique and lessen the loss of my proteins..
Any ideas?
Some initial questions: Lane i is the flow through, right? Are lanes 2 and 3 the elution? So which lane shows the load solution? In other words, do you have a gel which shows the whole story of the process where there's a lane for each process step that would include the load onto the dextrose column, the dextrose column elution, post TEV protease before dialysis, and then the load solution for the ion exchanger? Then, do you have gel lanes to demonstrate material balance where evderyting that went onto the column is seen coming off the column? Which lane is the strip of the column where the column is regenerated? There are a number of possibilities but with more information the explanation can be narrowed down.
A possible problem with your approach was that by adjusting the pH of the buffer to match the pI of the protein, you may have created a situation that encouraged precipitation of the protein. At the pI, the net charge of a protein is zero, but there may still be charges on the surface that allow it to bind to an ion exchange resin. This could allow the protein to become concentrated enough to precipitate in the column. (By the way, Tris is not very useful as a buffer at pH 6.4 because its pKa is 8.1.).
A smple way to remove the cleaved MBP is to run the sample over the dextrose affinity column again. The MBP will bind and the other protein will pass through.
I agree with Adam and would suggest the easy to do option of rerunning the protein after TEV treatment though the dextrose affinity column.This is what I do with my MBP-tagged protein and it works just fine. After running through the dextrose column i do a size exclusion chromatography step to polish the purity a bit more.
In your case, the 47 and 42 kDa are very close to make judgement based on SDS PAGE unless you are running a gradient gel. I hope you have run a SDS PAGE of Pre and Post cleaved protein.
Hi everyone, Thanks for all your suggestions!
Grant, lanes 1,2,3 are flowthrough. Lanes 4-11 are the elution. I did not runt he load solution sample on the gel.. Thank you for your suggestion; I will run a gel with all the lanes (uninduced, induced, post dextrose purification, post dialysis, post TEV cleavage, post second dextrose purification)
Alejandro, I've only MS bands 3 and 7. Both results returned saying that they're MBP. I'm thinking that my proteins may have precipitated in the process..
Adam and Venu, I'm testing out a new buffer in where I'm using HEPES instead of Tris and at pH 7. This time I will not be doing anionic exchange but a second run through the dextrose affinity column. I'll see what I get. Thanks again for your suggestions!
Hi,
I've tried using the new buffer and ran my sample under a second run of dextrose column. I've loaded a total of 1mg of my protein. It probably was diluted under the second run.. What I expected to see on my gel was the protein of my interest at the flowthrough fraction and the tag eluted with the maltose. I did see my tag eluted however there were no bands corresponding to the size of my protein. If I had an antibody towards my protein I wouldve attempt a western blot but currently there's not an antibody for it. any thoughts?
Lanes: 1 (post 1st dextrose purification)
2 (post dialysis)
3 & 4 (flowthrough from 2nd dextrose purification)
5-8 (wash with buffer A)
9-14 (elution with buffer B)
The molecular weight of the main band is larger than the 47 kDa protein or MBP, indicating that cleavage did not occur. This could be because of aggregation or dimerization of the fusion protein during dialysis, which might explain the appearance of the even higher molecular weight band. The full fusion protein appears to have bound to the column, but was not successfully eluted by maltose. Perhaps you need to use a higher concentration.
This is what I suspected could happen (yet, still no proof) and why I suggested running your gel samples both reduced and non-reduced as well as running a sample of the column strip (material balance). A column strip is not a high dextrose or high salt elution but to run something like 6M guanidine HCl or urea with maybe 0.5 M NaOH to be sure all protein is removed from the resin. Would you be surprised if those higher order bands collapsed to monomer mass upon reduction?
Also it seems the monomer may not be recovering well but you don't know until you do material balance which will imply a lot about what could be occurring. Covalent aggregates, if you prove their presence with reduced gels, indicate that the target moiety of the fusion is not folding well so that both aggregates and monomer may be mis-folded, albeit the MBP may be fine. Mis-folding can definitely obscure your TEV site; no cleavage with the site buried . Have you done a basic experiment of incubating the pre-dextrose column fusion molecule with maybe a 1:25 ratio of protease in solution (not on-column) and then running the gel reduced vs non-reduced (you might also pick up the condition of the protease)? I assume this was an E. coli expression system and so no glycosylation? I assume this is a eukaryotic protein with cysteines?
Another possibility: Etch virus protease is a cysteine protease but is there any trace of reducing agent and EDTA in your cleavage reactions (didn't see in your description)? TEV protease can also auto-inactivate but I believe there are versions that don't do this. If your protein needs to form disulfides for correct folding then you have to consider the order of folding in conjunction with a reductive cleavage condition. Run the process reduced and then fold? If is folds firstly with disulfide formation and then requires partially reductive cleavage conditions then what might happen?
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